Protocol Guide: Immunofluorescent Staining of Whole-Mount Organoids using Antibodies

Organoids have complex 3-dimensional structures and are embedded in ECM hydrogels making characterization difficult. Immunofluorescent (IF) staining of organoids with antibodies can be performed to visualize molecular markers of cellular behavior, proliferation, differentiation, cell health and identity. Whole-mount immunostaining is intended for small pieces of tissue without sectioning and methods are very similar to immunocytochemistry (ICC) or immunohistochemistry (IHC) staining of cryosections. Whole-mount staining of organoids can be used for antibody based characterization. In this organoid staining protocol we detail methods to fix, permeabilize, and stain whole-mount organoids for analysis by immunofluorescent confocal microscopy.

Organoid Staining Protocol

1. Remove media from organoid culture and gently wash 2X with 1X PBS (D8537).
2. Fix 30-40 organoid Matrigel domes in a 10 cm dish with 15-20 mL of 4% PFA (1.00496) for 30-60 minutes at room temperature.
   Note: Matrigel will partially dissolved when fixed in PFA.
3. Swirl the dish occasionally to detach Matrigel domes and to release organoids from Matrigel.
   Note: not all organoids will be released from Matrigel.
4. Pour the 15-20 mL fixative from the 10 cm dish into a 50 mL conical tube and allows the organoids to settle at the bottom of the tube by gravity (~10-15 minutes).
5. After the organoids have settled at the bottom of the 50 mL conical tube by gravity, carefully aspirate the fixative trying to avoid aspirating the organoid pellet in the process.
6. Add 15-20 mL of 1X PBS (D8537) to the dish from step 4 and let it sit for 15-20 minutes at room temperature.
7. When the 15-20 minutes is up, swirl the dish to detach the Matrigel domes further and pour the 1X PBS (D8537) into the 50 mL conical tube containing the organoid pellet from step 5.
8. Repeat steps 5-7 three more times.
9. Aspirate the solution without harming the organoid pellet and add 5 mL of 1X PBS (D8537) by dispensing on the side of the 50 mL conical tube containing the organoid pellet and swirl the tube to resuspend the organoid pellet.
   Note: do not pipet up and down with a pipet. This will break most of the organoids.
10. Directly pour the 5 mL of 1X PBS (D8537) containing the organoid clumps into the 10 cm dish containing the undetached organoid Matrigel domes.
11. Add another 5 mL of 1X PBS (D8537) into the 50 mL conical tube to collect as much of the organoid clumps as possible and directly pour it in the 10 cm dish with the organoids.
12. If not using immediately, seal the 10 cm dish with parafilm and store in the fridge at 4 °C for up to 1 month.
13. When ready to perform immunofluorescent staining, remove the 10 cm dish containing the fixed organoids from the fridge and observe under a dissecting microscope.
14. Cut the tip of a 1 mL pipet tip so that the barrel will be large enough to suck up the organoids without shearing or breaking them.
15. Transfer the organoids (1-4) into a 8-well chamber slide and remove residual 1X PBS (D8537) with a P-200 pipet. Avoid sucking up the organoids and shear them through the uncut P-200 tip.
16. Permeabilize the fixed organoids with 0.5 mL of Blocking Buffer (5% horse serum + 0.5% Triton X-100 in 1X PBS) overnight at 4 °C or 2-4 hours at room temperature. Note: it’s recommended to use the serum from the same species as the host of the secondary antibody.
17. Remove Blocking Buffer from the chamber slide containing the organoids with a P-200 pipet. Avoid pipetting up the organoids through the P-200 tip.
18. Prepare primary antibodies (300-500 µL) or direct conjugated antibodies (300-500 µL) in Blocking Buffer.
19. Add diluted antibodies into the appropriately labeled well containing the organoids.
20. Allow to incubate at 4 °C overnight.
21. On the next day, wash 3X with 1X PBS (D8537) for 10-15 minutes each wash. Note: If using direct conjugated antibodies samples are ready to be imaged on a confocal microscope.
22. If using unconjugated primary antibodies, prepare secondary antibodies (300-500 µL) in Blocking Buffer and to the appropriately labeled samples.
23. Allows to stain with secondary antibody overnight at 4 °C.
24. On the next day, wash 3X with 1X PBS (D8537) for 10-15 minutes each wash.
25. Remove Blocking Buffer from each well containing the organoids with a P-200 pipet. Avoid pipetting up the organoids through the P-200 tip.
26. Prepare nuclear staining by added DAPI (D9542) at 5 µg/mL in 1X PBS (300-500 µL per sample).
27. Add DAPI staining solution to each sample and allows to incubate at room temperature for 15-20 minutes.
28. Wash 3X with 1X PBS (D8537) for 10-15 minutes each wash.
29. Samples are ready to be imaged on a confocal microscope.